altering sphingolipid metabolism in saccharomyces ...cycling the plasma membrane at synaptic...

EUKARYOTIC CELL, May 2009, p. 779–789 Vol. 8, No. 51535-9778/09/$08.00�0 doi:10.1128/EC.00037-09Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Altering Sphingolipid Metabolism in Saccharomyces cerevisiae CellsLacking the Amphiphysin Ortholog Rvs161 Reinitiates Sugar

Transporter Endocytosis�

Jeanelle Morgan,2†‡ Paula McCourt,1,2† Lauren Rankin,1 Evelyn Swain,2§Lyndi M. Rice,1,2 and Joseph T. Nickels, Jr.1,2*

Pharmacogenomics Division, Medical Diagnostics Laboratories, L.L.C., Hamilton, New Jersey 08690,1 and Department ofBiochemistry and Molecular Biology, Drexel University College of Medicine, Philadelphia, Pennsylvania 191022

Received 26 January 2009/Accepted 6 March 2009

Amphiphysins are proteins thought to be involved in synaptic vesicle endocytosis. Amphiphysins share acommon BAR domain, which can sense and/or bend membranes, and this function is believed to be essentialfor endocytosis. Saccharomyces cerevisiae cells lacking the amphiphysin ortholog Rvs161 are inviable whenstarved for glucose. Altering sphingolipid levels in rvs161 cells remediates this defect, but how lipid changessuppress remains to be elucidated. Here, we show that the sugar starvation-induced death of rvs161 cellsextends to other fermentable sugar carbon sources, and the loss of sphingolipid metabolism suppresses thesedefects. In all cases, rvs161 cells respond to the starvation signal, elicit the appropriate transcriptionalresponse, and properly localize the requisite sugar transporter(s). However, Rvs161 is required for transporterendocytosis. rvs161 cells accumulate transporters at the plasma membrane under conditions normally result-ing in their endocytosis and degradation. Transporter endocytosis requires the endocytosis (endo) domain ofRvs161. Altering sphingolipid metabolism by deleting the very-long-chain fatty acid elongase SUR4 reinitiatestransporter endocytosis in rvs161 and rvs161 endo� cells. The sphingolipid-dependent reinitiation of endocy-tosis requires the ubiquitin-regulating factors Doa1, Doa4, and Rsp5. In the case of Doa1, the phospholipaseA2 family ubiquitin binding motif is dispensable. Moreover, the conserved AAA-ATPase Cdc48 and its acces-sory proteins Shp1 and Ufd1 are required. Finally, rvs161 cells accumulate monoubiquitin, and this defect isremediated by the loss of SUR4. These results show that defects in sphingolipid metabolism result in thereinitiation of ubiquitin-dependent sugar transporter endocytosis and suggest that this event is necessary forsuppressing the nutrient starvation-induced death of rvs161 cells.

The budding yeast Saccharomyces cerevisiae gene RVS161/END6 encodes a helical protein of 265 amino acids that is amember of the N-BAR (for Bin, amphiphysin, Rvs) family ofproteins (61). Rvs161 regulates cell polarity (20), actin cy-toskeleton polarization (69), endocytosis (50), and secretoryvesicle trafficking (7, 25). rvs161 mutant cells die during sta-tionary phase, have mating defects, are sensitive to high con-centrations of NaCl, have endocytosis and actin defects, andare unable to grow on nonfermentable carbon sources (8, 13,15, 50, 63, 69). Mutational studies have revealed two function-ally independent Rvs161 domains: an NH2-terminal/BAR do-main involved in endocytosis and actin organization and aCOOH-terminal domain required for cell fusion during hap-loid cell mating (8).

The N-BAR family of proteins is constantly growing andincludes yeast Rvs161 and Rvs167; human BIN1, BIN2, andBIN3; hob1� and hob3� from Saccharomyces pombe; murine

Alp1; and human amphiphysins, which are a family of multido-main proteins that are involved in the late steps of clathrin-coated vesicle scission (24, 30, 58, 61). Two genes encodeamphiphysin in humans, one expressed in the brain (am-phiphysin I) and a second (amphiphysin II) with a broad tissuedistribution and a wide array of alternatively spliced isoforms.Both have an N-terminal BAR domain and a C-terminal SH3domain. The BAR domain interacts with phospholipids, whichinduces membrane curvature and tubulation, whereas the dis-tal SH3 domain associates with proteins having proline-basedmotifs. Human amphiphysin has been implicated in endocyto-sis due to its interaction with dynamin and because of itshomology to Rvs161 and Rvs167. It may be important in re-cycling the plasma membrane at synaptic terminals (14, 28,42, 68).

The crystal structures of several N-BAR domains have beensolved (9, 45, 58, 71, 72). The domain is present in proteins thatare critical for the recycling of synaptic vesicles and T-tubuleformation in muscle, such as amphiphysins, endophilin, nadrin,beta-centaurins, arfaptin, and oligophrenins. Peter et al. (58)have shown that an N-BAR domain (the BAR domain plus anadjacent amphipathic helix) is capable of inducing three-dimensional membrane curvature. Some BAR proteins haveadditional interactions with lipids through pleckstrin homologyor phox homology domains. These domains may target a pro-tein to a specific membrane compartment, while the BARdomain simultaneously detects or initiates membrane curva-

* Corresponding author. Mailing address: Pharmacogenomics Divi-sion, Medical Diagnostics Laboratories, L.L.C., 2439 Kuser Road,Hamilton, NJ 08690. Phone: (609) 570-1046. Fax: (609) 570-1060.E-mail: [email protected].

† These authors contributed equally to the manuscript.‡ Present address: Department of Food Science, Rutgers University,

New Brunswick, NJ 08901.§ Present address: Department of Chemistry, Newberry College,

Newberry, SC 29108.� Published ahead of print on 13 March 2009.

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ture. There are two other BAR domains, F-BAR and I-BAR,which bind membranes and in some cases induce membranetubulation (10, 35).

rvs161 and rvs167 mutants have common phenotypes (2, 7,63, 69), and Rvs161 and Rvs167 physically interact (70). How-ever, they have distinct nonoverlapping cell functions andphysical interactions with other proteins (8, 26). The BARdomains of each cannot be interchanged, and the overexpres-sion of RVS161 or RVS167 cannot cross-suppress the other’sphenotype (2, 70). Defects of rvs cells, including salt sensitivity,cell death during starvation, and the lack of growth on nonfer-mentable carbon sources, are suppressed by mutations alteringthe sphingolipid composition. SUR1, SUR2, SUR4, and IPT1encode inositolphosphorylceramide mannosyltransferase,long-chain-base (LCB) C4-hydroxylase, very-long-chain fattyacid elongase, and mannose diinositolphosphorylceramide syn-thase, respectively, and are required for the biosynthesis ofyeast complex sphingolipids (Fig. 1) (17, 19). Recessive muta-tions in these genes alter the amount and composition of com-plex sphingolipids (4, 18, 29, 51) and suppress rvs defects (1,15). Suppression may function through remediating the actindepolarization/repolarization defects seen in mutant cells intimes of stress (1). However, rvs161 sur4 and rvs167 sur4 cellshave steady-state actin defects when starved for glucose (26).Thus, the molecular basis of suppression is complex and re-mains to be uncovered.

The HXT family of proteins are mammalian-facilitated glu-cose transporter (GLUT) orthologs (44, 56, 76). S. cerevisiaehas 20 genes encoding proteins similar to hexose transporters(56). Most are bona fide transporters, such as Hxt1 to Hxt17,while others, such as Snf3 and Rgt2, are glucose sensors (54).Snf3 and Rgt2 sense extracellular glucose concentrations andinitiate a transcriptional signaling cascade (53, 54), resulting inthe expression of high-affinity (Hxt2 and Hxt4) or low-affinity(Hxt3 and Hxt1) transporters (55, 60). What is known aboutthe stability and degradation of glucose transporters is thatunder specific conditions, components of the high- and low-affinity glucose uptake system are inactivated (6). Studies haveexamined the stability and degradation pathway for Hxt6 andHxt7 (40). It is generally thought that glucose transporters areinternalized via endocytosis and subsequently degraded.

rvs161 cells die under conditions of glucose starvation. Here,we show that they harbor starvation defects on other ferment-able carbon sources and are unable to thrive when galactose,maltose, or melibiose is the available carbon source. Mutantcells can sense a glucose starvation signal, derepress glucose-repressed genes, initiate Snf3- and Rgt2-dependent HXT tran-scription, and properly localize high- and low-affinity glucosetransporters. They also express and properly localize the Gal2galactose and Mal61 maltose permeases. However, rvs161 cellsare unable to endocytose and degrade these sugar transporters.The loss of function of SUR4 suppresses all carbon sourcegrowth defects we observed and restores sugar transporterendocytosis and degradation. Doa1, Doa4, and Rsp5 arerequired for sur4-dependent suppression and for transporterendocytosis and degradation, as is the conserved AAA-ATPase Cdc48 and its accessory factors, Shp1 and Ufd1.

MATERIALS AND METHODS

Media and miscellaneous microbial techniques. Yeast strains were grown inYEP (1% yeast extract, 2% Bacto-peptone) containing the indicated concentra-tions of the indicated carbon source in YPG (1% yeast extract, 2% Bacto-peptone, 3% glycerol) or in synthetic minimal medium containing 0.67% yeastnitrogen base (Difco) supplemented with the appropriate amino acids and ade-nine. Yeast transformations were performed using the procedure described pre-viously (34). For the routine propagation of plasmids, Escherichia coli XL1-Bluecells were used and grown in Luria broth supplemented with ampicillin (200mg/ml).

Strain and plasmid construction. The yeast strains used are derived fromW303 (YJN17) (MATa ura3-52 leu2 his3 lys2 ade2). hxt1::HXT1-GFP::TRP1,hxt2::HXT2-GFP::TRP1, hxt4::HXT4-GFP::TRP1, gal2::GAL2-GFP::TRP1, andmal61::MAL61-GFP::TRP1 alleles were generated as described previously (43)using the pFA6a-GFP(S65T)-TRP1 module. cdc48::kanr ura3-52::cdc48-3::URA3and ufd1::kanr ura3-52::ufd1-1::URA3 strains were generated using the diploidstrain YJN1 (MATa/� ura3-52/ura3-52 his3/his3 leu2/leu2 TRP1/trp1 LYS2/lys2ade2/ade2), as the deletion of CDC48 or UFD1 is lethal in haploid strains. First,cdc48::kanr and ufd1::kanr alleles were synthesized by the PCR amplification ofthe cdc48::kanr or ufd1::kanr allele from heterozygous CDC48/cdc48::kanr andUFD1/ufd1::kanr strains (Research Genetics), respectively. These alleles weretransformed into YJN1, and integrants were selected on yeast extract-peptone-dextrose (YEPD) plates containing 250 �g/ml G418. Proper integration wasdetermined by PCR. YIp-cdc48-3 and YIp-ufd1-1 were digested with StuI andintegrated at the ura3-52 locus of the CDC48/cdc48::kanr and UFD1/ufd1::kanr

strains, respectively. Cells were sporulated, and haploid cdc48::kanr ura3::cdc48-3::URA3 and ufd1::kanr ura3-52::ufd1-1::URA3 strains were obtained byprototrophic amino acid and temperature-sensitive selections. The cdc48-3 andufd1-1 alleles used to construct YIp-cdc48-3 and YIp-ufd1-1 were generated bythe PCR amplification of YJN3151 and YJN3158, respectively. doa1::HIS3,doa1::doa1�C::TRP1, and doa1::doa1F417D F434D::TRP1 alleles were generatedby PCR amplification using YJN3222, YJN3233, and YJN3225, respectively.shp1::kanr strains were generated by the PCR amplification of the shp1::kanr

allele from a haploid shp1::kanr strain (Open Biosystems, Huntsville, AL) andsubsequent to the transformation of the PCR product into YJN17 and selectionon G418. pGAL-rsp5-1 was used to express the dominant-negative rsp5-1 allele.

Total RNA isolation. All solutions were prepared with diethyl pyrocarbonate-treated water. Cells were harvested, centrifuged, and pelleted for 30 s. Cellpellets were resuspended in 200 �l of YRL buffer (200 mM Tris, pH 7.5,containing 500 mM NaCl, 10 mM EDTA, 1% sodium dodecyl sulfate [SDS]) and200 �l PCIAA (phenol, chloroform, isoamyl alcohol). Two hundred microlitersof nitric acid-washed beads was added, and cells were vortexed for 2.5 min. Threehundred microliters of YRL buffer and 200 �l PCIAA were added, and cellswere vortexed for 2.5 min. Cells were centrifuged for 5 min, and the resultingclear lysate was removed and added to 400 �l PCIAA, vortexed for 2.5 min, andcentrifuged for 5 min. The resulting aqueous layer was added to 500 �l of 100%ethanol, and total RNA was precipitated overnight at �20°C. A total RNA pelletwas obtained by centrifugation at 13,000 rpm for 15 min at 4°C, washed twicewith 70% ethanol, vacuum dried, and resuspended in water. RNA was stable at�20°C for several weeks.

FIG. 1. Sphingolipid biosynthetic pathway in S. cerevisiae. Genesare in italics. FA, fatty acid; VLCFA, very long chain fatty acids.

780 MORGAN ET AL. EUKARYOT. CELL

Northern analysis. Total RNA was resolved using 6% formaldehyde agarosegel electrophoresis. Total RNA (20 �g) in loading buffer (20 mM morpho-linepropanesulfonic acid, pH 7.0, containing 10 mM sodium acetate, 2 mMEDTA, 45% formamide, 6% formaldehyde, 1% ethidium bromide, 0.003% bro-mophenol blue, 0.03% xylene cyanol FF, 1.5% Ficoll) was analyzed. RNA wasblotted onto Hybond-N nitrocellulose (Amersham, Arlington Heights, IL) over-night at room temperature using 10� SSC (1� SSC is 0.15 M NaCl plus 0.015 Msodium citrate). Churches buffer (10 mM sodium-phosphate buffer, 1 mMEDTA, 1% bovine serum albumin, 7% SDS) was used for all hybridizationprocedures. Hybridization was performed overnight at 65°C. Gel-purified radio-labeled probes were boiled in 200 �l salmon sperm DNA prior to use. Afterhybridization, blots were washed twice in 2� SSC at room temperature, twice in2� SSC–0.5% SDS at 65°C, and twice in 0.1� SSC at room temperature. Geneexpression was determined by autoradiography using Kodak X-OMAT film. Thespecificity of each HXT probe was checked by Northern analysis using hxt1, hxt2,hxt3, and hxt4 strains. U2 expression was used as a loading control.

Fluorescence microscopy. Cultures were grown to exponential phase in YEPcontaining 6% glucose. Cells then were shifted to YEP containing the indi-cated concentrations of glucose for the specified time. Hxt1-green fluorescentprotein (Hxt1-GFP), Hxt2-GFP, and Hxt4-GFP localization was visualizedusing a Leica DRBME fluorescence microscope, fluorescein isothiocyanateoptics, and a PlanAPO �100 objective. Images were obtained using OpenLabs software (version 2.1). Final fluorescence images were generated usingAdobe Photoshop (version 7.0).

Western analysis of Hxt1-GFP and Hxt2-GFP stability. Cultures were grownto exponential phase (optical density at 600 nm of 0.5 to 1.0) in YEPD and thenshifted to the appropriate medium to regulate HXT expression and proteinsynthesis. Twenty-milliliter cultures were collected at various times, and a totalcell protein extract was obtained. Briefly, cells were pelleted, washed once withdistilled water, and lysed in buffer A [200 mM Tris-HCl, pH 7.9, containing 390mM (NH4)2SO4, 10 mM MgSO4, 20% (vol/vol) glycerol, 1 mM EDTA] usingglass beads. Lysis buffer also contained 15 mM mercaptoethanol, 1 mM AEBSF[4-(2-aminoethyl)-benzenesulfonyl fluoride], 5 �g/ml pepstatin, 5 �g/ml leupep-tin, and 10 �l of Sigma protease inhibitor solution (Sigma Chemicals, St. Louis,MO). Five hundred micrograms of total cell protein extract was resuspended inLaemmli buffer (12), and proteins were resolved using 10% SDS–polyacrylamidegel electrophoresis (PAGE) and transferred to nitrocellulose membranes.

All steps for Western analysis were performed at room temperature. Mem-branes were blocked for 1 h with 5% nonfat dry milk in buffer C (10 mMTris-HCl, pH 7.4, containing 150 mM NaCl and 0.05% Tween-20). Incubationswith primary and secondary antibodies were performed for 1 h in buffer C.Membranes were washed six times after antibody incubations with buffer C. Blotswere incubated with mouse anti-GFP monoclonal antibodies (1:2,000 dilution)(Clontech, Mountain View, CA) and anti-mouse horseradish peroxidase (HRP)-conjugated monoclonal antibodies (1:5,000 dilution) (Amersham Corp., Arling-ton Heights, IL). Anti-actin polyclonal antibodies (1:1,000) (Santa Cruz Biotech-nology, Santa Cruz, CA) were used to determine protein loading. Hxt1-GFP andHxt2-GFP were detected using chemiluminescence and autoradiography. Anti-Gal2 and anti-Mal61 polyclonal antibodies were used at a 1:500 dilution.

Detection of extracellular invertase protein levels. Suc2 invertase expressionwas induced as described previously (57). Secreted Suc2 was determined byWestern analysis using anti-Suc2 polyclonal antibodies at a 1:500 dilution.

Western analysis of total cellular ubiquitin. Total cell extracts were obtainedusing a modified procedure from Mullally et al. (49). Yeast cells grown to late logphase (optical density at 600 nm of 2.0) in synthetic complete medium containing2% glucose were pelleted and resuspended in whole-cell lysis buffer (50 mMTris-HCl, pH 7.4, 150 mM NaCl, 1 mM dithiothreitol, 0.05% Triton X-100)containing complete protease inhibitors (Roche Applied Science, Indianapolis,IN). Cells then were lysed with glass beads by using four cycles of vortexing for1 min and incubation on ice for 1 min. Cellular extracts were collected aftercentrifugation, and samples were boiled in SDS loading buffer. Proteins wereresolved by SDS–18% PAGE and subsequently transferred to a nitrocellulosemembrane that was boiled in distilled deionized water for 10 min prior toblocking. Membranes were treated as described for the detection of Hxt2 andHxt4. The primary antibody was monoclonal mouse anti-ubiquitin (clone P4D1;1:500 dilution; Santa Cruz Biotechnology, Santa Cruz, CA), and the secondaryantibody was polyclonal goat anti-mouse immunoglobulin G HRP-conjugatedantibody (1:1,000 dilution; GE-Amersham, Piscataway, NJ). Proteins were de-tected using ECL chemiluminescence (GE-Amersham, Piscataway, NJ). For aloading control, actin protein was detected using monoclonal rabbit anti-�-actinprimary antibody (clone H-300; 1:500 dilution; Santa Cruz Biotechnology) andpolyclonal goat anti-rabbit immunoglobulin G HRP-conjugated antibody (1:1,000 dilution; GE-Amersham). Densitometry was performed using a Bio-Rad

GS-800 densitometer and Quantity One software (4.6.6). All relative densitom-etry values were normalized to total actin levels. The values in Fig. 9 are theaverages from three experiments � standard deviations.

RESULTS

rvs161 cells harbor pleiotrophic carbon source growth de-fects that can be suppressed by altering sphingolipid levels.Cells lacking RVS161 cannot grow under conditions of glucosestarvation (Fig. 2A). As previously described (15), this starva-tion defect could be suppressed by altering sphingolipid levelsthrough the loss of SUR4. We asked whether starvation-in-duced death extended to other fermentable carbon sources.rvs161 cells were unable to grow when starved for fructose,sucrose, or raffinose (Fig. 2A). Interestingly, mutant cells alsowere incapable of growing on 2% galactose, 2% maltose, or2% melibiose (Fig. 2B). Deleting SUR4 remediated all carbonsource defects we observed (Fig. 2). To uncover the molecularbasis for these carbon source defects and their sphingolipid-dependent suppression, we asked whether rvs161 cells couldsense changes in glucose levels or distinguish what carbonsource they were grown on and regulate gene transcriptionaccordingly.

rvs161 cells are able to derepress glucose-repressed genes aswell as activate carbon source-dependent transcription. Glu-cose represses the expression of genes that are dispensableunder rich growth conditions (e.g., GAL, SUC, and genes en-coding cytochromes and tricarboxylic acid cycle enzymes) (54).When glucose levels are depleted, the expression of these andother genes are induced or derepressed, and this transcrip-tional response is required for sustained growth. We askedwhether rvs161 cells were capable of derepressing glucose-repressed genes. As a model for glucose derepression, we ex-amined the expression levels of the invertase gene SUC2.

Cells were grown in YEPD and shifted to glycerol-contain-ing medium to induce SUC2 expression. All strains tested wereable to derepress and induce the expression of SUC2 (Fig. 3A).We also tested if mutant cells could induce SUC2 expression inraffinose-grown cultures. We reasoned that raffinose utilization(2% raffinose) through invertase-dependent hydrolysis wouldmore closely mimic low-glucose conditions while being lessdeleterious to cells than examining expression in glucose star-

FIG. 2. rvs161 cells harbor pleiotrophic carbon source starvationgrowth defects. Tenfold serial dilutions were spotted onto YEP platescontaining the indicated sugar carbon source concentration. The initialcell density was 1 � 105 cells/ml. Cells were grown for 2 days at 30°C.WT, wild type; Glu, glucose; Fru, fructose; Suc, sucrose; Gal, galac-tose; Mal, maltose; Mel, melibiose.

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vation medium (0.05% glucose), a condition that kills rvs161cells. The induction of SUC2 expression in raffinose has beendemonstrated previously (57). SUC2 expression was induced inall strains tested and was sustained to similar levels (Fig. 3B).rvs161 cells were incapable of growing on galactose and mal-tose (Fig. 2B). Thus, we examined the galactose- and maltose-induced expression of the GAL2 and MAL61 permease genes,respectively. We found no differences in expression betweenwild-type and rvs161 cells (Fig. 3C). Finally, the sucrose-in-

duced extracellular excretion of Suc2 invertase that is requiredfor raffinose hydrolysis was normal in rvs161 mutants (data notshown). Based on these results, we conclude that rvs161 cellsdo not harbor defects in derepressing glucose-repressed genes,in their transcriptional response to growth on alternative fer-mentable carbon sources, or in secreting enzymes that arerequired to grow on various carbon sources.

rvs161 cells activate the Snf3- and Rgt2-dependent expres-sion of HXT genes. Glucose utilization in S. cerevisiae beginswith the transport of glucose into the cell by specific high- andlow-affinity glucose transporters. Hxt2 and Hxt4 are high-affinity glucose transporters and are expressed when glucoselevels are low, while the expression of the low-affinity glucosetransporter Hxt1 is induced in high glucose concentrations.The expression of HXT3 is moderately induced at all glucoseconcentrations (55). The glucose-dependent expression ofHXT transporters is initiated and terminated by the Snf3 andRgt2 glucose sensors. We asked whether the lack of properHXT expression contributed in any way to starvation-induceddeath by asking if rvs161 cells were capable of initiating andterminating Snf3- and Rgt2-dependent HXT expression. Glu-cose concentration-dependent HXT expression levels were de-termined using Northern analysis. The expression patterns ofHXT1 to HXT4 in wild-type and rvs161 cells grown undervarious glucose concentrations were identical (Fig. 3D). Thus,rvs cells are able to initiate Snf3- and Rgt2-dependent tran-scriptional signaling, resulting in the proper expression of HXTgenes.

rvs161 cells are defective in the endocytosis and degradationof multiple sugar transporters. Normal HXT gene expressionobserved in rvs161 cells prompted us to determine whether Hxttransporter mislocalization and/or protein instability contrib-uted to starvation-induced death. rvs161 mutants do accumu-late vesicles at the cytoplasmic side of the plasma membrane(7, 25). We constructed strains harboring an endogenous GFP-tagged allele of HXT2. Various glucose levels regulated HXT2gene expression, and fluorescence microscopy was used to vi-sualize localization.

As expected based on our expression data, plasma mem-brane-associated Hxt2 was not seen in wild-type cells grown inrich medium (data not shown). It localized to the plasma mem-brane after cells were shifted to starvation media for 2 h(0.05% Glu), and it disappeared after a shift back to richmedium for 5 h (Fig. 4). Thus, wild-type cells properly local-ized Hxt2 in response to changes in glucose levels. We foundthat Rvs161 was dispensable for the plasma membrane-asso-ciated localization of Hxt2 but was absolutely required for itsdisappearance (Fig. 4). Hxt2-GFP levels accumulated at theplasma membrane in mutant cells even after being shifted torich medium for 5 h and persisted for up to 16 h. Importantly,we could reinitiate the loss of Hxt2 at the plasma membrane ifwe deleted SUR4 in mutant cells (Fig. 4). Interestingly, cellslacking SUR4 showed a fluorescent fragmented vacuolar mor-phology rather than the large single fluorescent organelle seenin cells with normal Sur4 function. This has been observedpreviously (39).

To determine whether the disappearance of Hxtp from theplasma membrane was representative of their endocytosis anddegradation, the kinetics of glucose transporter degradationwere determined using Western analysis. The high-affinity glu-

FIG. 3. Transcriptional responses of rvs161 cells to growth on mul-tiple carbon sources and carbon source starvation are intact. In allcases, cells were grown to exponential phase in YEPD at 30°C prior toshifting cells to the various carbon sources. Gene expression levelswere determined by Northern analysis. (A) Cells were shifted to me-dium containing 2% glycerol–0.05% glucose. (B) Cells were shifted tomedium containing 2% raffinose. Cells were shifted to medium con-taining 2% galactose for GAL2 expression (C) or 2% maltose forMAL61 expression (D). (E) Lane 1, cells were shifted to 2% glucose,and HXT expression levels were determined after 2 h; lane 2, cells wereshifted to 0.05% glucose, and HXT expression levels were determinedafter 2 h; lane 3, cells were shifted back to 2% glucose and HXTexpression levels were determined after 2 h. U2 expression was used asa loading control.


cose transporters Hxt2 and Hxt4 were examined first. Hxt2 wasdetected in wild-type cells after 2 h in 0.05% glucose-contain-ing medium and was almost completely endocytosed and de-graded after a 3-h shift to rich medium (Fig. 5A). sur4 mutantsshowed similar kinetics (Fig. 5A). In contrast, rvs161 cells ac-cumulated Hxt2 (Fig. 5A), while degradation was restored ifwe deleted SUR4 (Fig. 5A). Identical results were obtainedwhen examining Hxt4 degradation (data not shown).

The endocytosis and degradation of low-affinity glucosetransporters also were defective in rvs161 cells, as evidenced bythe aberrant accumulation of Hxt1. We found that Hxt1 wasplasma membrane localized in wild-type cells grown in me-dium containing 2% glucose (Fig. 5B). After 1 h of beingshifted to medium containing 0.05% glucose, wild-type cellscompletely endocytosed and degraded this transporter (Fig.5B). rvs161 cells accumulated Hxt1 under these same condi-tions (Fig. 5B). The loss of SUR4 in rvs161 cells restored Hxt1endocytosis and degradation levels to those seen in sur4 cells(Fig. 5B). Fluorescence microscopy showed that Hxt1-GFPlocalized to the plasma membrane in rvs161 cells (data notshown).

rvs161 cells also were defective in endocytosing and degrad-

ing the galactose permease Gal2 (Fig. 5C). Once again, delet-ing SUR4 restored endocytosis (Fig. 5C). Identical results wereobtained by examining the endocytosis of the maltose per-mease Mal61 (data not shown). Both Gal2-GFP and Mal61-GFP localized to the plasma membrane in rvs161 cells (datanot shown). Based on our results, we conclude that rvs161 cellsharbor a general defect in sugar transporter endocytosis that canbe remediated by altering sphingolipid metabolism through theloss of SUR4.

The endocytosis domain of Rvs161 is required for growthunder glucose starvation and for the endocytosis of glucosetransporters. The fact that Rvs161 is required for growth onmultiple carbon sources and for the endocytosis of multiplesugar transporters prompted us to determine the Rvs161 do-

FIG. 4. Aberrant plasma membrane accumulation of Hxt2 inrvs161 cells can be suppressed by the loss of SUR4. Cells were grownto exponential phase at 30°C in YEPD and then sequentially shifted tomedium containing the indicated sugar concentrations for the indi-cated times. Hxt2-GFP localization was visualized by live-cell fluores-cence microscopy. WT, wild type; Glu, glucose; hr, hours.

FIG. 5. Sugar transporter endocytosis defects of rvs161 cells can besuppressed by the loss of SUR4. Cells were grown to exponential phaseat 30°C in YEPD (A and B) or YEP containing 2% galactose (C). Theythen were sequentially shifted to medium containing the indicatedsugar concentrations for the times indicated. Cell lysates were resolvedby SDS-PAGE, and sugar transporter levels were determined by West-ern analysis using anti-GFP polyclonal antibodies (A and B) or anti-Gal2 polyclonal antibodies (C). Act1 (actin) levels were used as aloading control. WT, wild type; Glu, glucose; hr, hours. (A) Hxt2-GFP;(B) Hxt1-GFP; (C) Gal2.


main(s) responsible for these functions. Brizzio et al. (8) iso-lated strains harboring recessive rvs161 alleles giving rise toeither endocytosis (End� Fus�) or cell fusion/mating (End�

Fus�) defects, thus delineating the functional domains ofRvs161. R35C, R113K, and P158S alleles cause endocytosisdefects, while A175P and P203Q are defective in cell fusion/mating based on several criteria (Fig. 6A) (8).

We tested whether the two cell fusion and three endocytosismutants (P158S, R35C, and R113K) could grow when starvedof glucose. Surprisingly, none of these mutants harbored de-fects at 30°C (data not shown). Only when we grew cells at37°C did we observe a phenotype (Fig. 6B). rvs161 cells har-boring P158S, R35C, or R113K (End� Fus�) alleles were un-able to grow. A175P cells (End� Fus�) grew as well as wild-type cells, but P203Q cells consistently showed an intermediatestarvation defect (Fig. 6B). We obtained similar results withthe carbon sources and various sugar starvation conditionstested in Fig. 2 (data not shown).

We next asked if the endo domain was required for glu-cose transporter endocytosis. We examined the localizationand endocytosis of Hxt2-GFP by using rvs161 R113K as arepresentative endo� mutant (Fig. 7). At 30°C, mutant cellslocalized, endocytosed (with a slight delay), and degradedHxt2-GFP like wild-type cells. However, at 37°C, rvs161R113K cells were incapable of endocytosing Hxt2-GFP fromthe plasma membrane. The loss of SUR4 suppressed thisdefect (data not shown). Cells harboring the A175P allelewere not defective in endocytosis (data not shown). Similarresults were observed for Hxt1-GFP, Gal2-GFP, and Mal61-GFP localization and degradation (data not shown). Thus,the endocytosis domain of Rvs161 is required for growth

under glucose starvation and for the endocytosis and deg-radation of multiple sugar transporters.

Reinitiation of Hxt2 endocytosis in rvs161 sur4 requiresDoa1, Doa4, and Rsp5 functions. We next asked if factorsrequired for ubiquitin-mediated endocytosis were needed forthe sur4-dependent reinitiation of sugar transporter endocyto-sis. To address this, we determined if Hxt2-GFP endocytosis inrvs161 sur4 cells required Doa1 (which regulates the cellularubiquitin concentration), Doa4 (ubiquitin hydrolase), and/orRsp5 (ubiquitin ligase). Fluorescence microscopy revealed arequirement for all three proteins. rvs161 sur4 doa1, rvs161 sur4doa4, and rvs161 sur4 cells harboring the dominant-negativersp5-1 allele all accumulated Hxt2-GFP under conditions ofhigh glucose growth (Fig. 8). Thus, the reinitiation of glucosetransporter endocytosis in rvs161 cells by the loss of SUR4requires several factors for ubiquitin-mediated endocytosis.

The Cdc48 binding domain of Doa1 is required for thereinitiation of glucose transporter endocytosis in rvs161 sur4cells. Doa1 contains a ubiquitin binding domain (PFU) and asecond carboxyl-terminal domain (PUL), which binds to theconserved AAA-ATPase Cdc48 (49). Both domains link ubiq-uitylated substrates to Cdc48 and are thought to be requiredfor Doa1 function (49, 62). We asked if one or both of these

FIG. 6. endo domain of Rvs161 is required for growth under con-ditions of glucose starvation. (A) Schematic of Rvs161 delineating thedomains required for endocytosis or cell fusion/mating, indicating themutations described by Brizzio et al. (8). (B) Tenfold serial dilutionswere spotted onto YEP plates containing the indicated sugar carbonsource concentration. The initial cell density was 1 � 105 cells/ml. Cellswere grown for 2 days at 37°C. WT, wild type; Glu, glucose.

FIG. 7. endo domain of Rvs161 is required for sugar transporterendocytosis. Cells were grown to exponential phase at 30°C in YEPD.The culture was split, and cells were incubated at 30 or 37°C in YEPcontaining the indicated sugar carbon source concentration for thetimes indicated. Hxt2-GFP localization was visualized by live-cell flu-orescence microscopy. WT, wild type; Glu, glucose; hr, hours.


domains were required for the sur4-dependent reinitiation ofsugar transporter endocytosis in rvs161 cells. We also deter-mined whether Cdc48 itself, and/or the Cdc48 accessory fac-tors Shp1 and Ufd1, also were needed. Shp1 and Ufd1 binddirectly to Cdc48 (46). Both Shp1 and Ufd1 bind ubiquitin.Shp1 is necessary for Cdc48 function in membrane fusion andproteosomal degradation, while Ufd1 facilitates endoplasmicreticulum (ER)-dependent degradation and the activation ofmembrane-associated transcription factors. We examined theglucose concentration-dependent appearance/disappearanceof Hxt2-GFP from the plasma membrane, and the data arepresented in Table 1.

The Cdc48 binding PUL domain of Doa1 (doa1�C allele)was required to reinitiate endocytosis by the loss of SUR4,while the PFU domain (doa1F417D F434D allele) was dispens-able. rvs161 sur4 doa1�C cells accumulated Hxt2 to the sameextent as rvs161 cells shifted from 0.05 to 6% glucose for 5 h,while the accumulation of this transporter in rvs161 sur4doa1F417D F434D cells was similar to that seen in rvs161 sur4suppressor cells (Table 1). Cdc48 itself also was required, asrvs161 sur4 cells harboring a cdc48-3 ts allele were incapable ofendocytosing Hxt2 at the restrictive temperature. In fact,rvs161 sur4 cdc48-3 cells showed a defect in endocytosis anddisplayed the characteristic elongated phenotype of cdc48 al-leles even at the permissive temperature (62). Both of theCdc48 binding factors, Shp1 and Ufd1, also were required.rvs161 sur4 cells deleted for SHP1 or harboring a temperature-sensitive ufd1-1 allele were defective in Hxt2 endocytosis (Ta-ble 1). Based on these results, we conclude that the sphingo-lipid-dependent reinitiation of sugar transporter endocytosis inrvs161 cells requires the function of multiple Cdc48 complexes.

Loss of SUR4 remediates the monoubiquitin hyperaccumu-lation defect of rvs161 cells. The gene products that are re-quired for the sur4-dependent suppression of rvs161 endocy-

tosis defects bind to and/or regulate ubiquitin levels.Moreover, our attempt to suppress the endocytosis defects ofrvs161 cells by overexpressing ubiquitin failed. Thus, we askedwhether rvs161 cells have an altered ubiquitin metabolism, andif so, does the loss of SUR4 suppress these defects. rvs161 cellshyperaccumulated monoubiquitin (1.3-fold), and deletingSUR4 in these cells did decrease the level of this ubiquitinspecies to nearly that seen in wild-type cells (80%) (Fig. 9).Interestingly, sur4 cells had a drastically lower level of mono-ubiquitin than the wild type, but their levels of high-molecular-weight conjugates were normal (Fig. 9). When we looked atthose mutations in rvs161 sur4 cells causing the loss of glucoseconcentration-dependent Hxt2 endocytosis (Table 1), the over-all observation was that they all altered ubiquitin metabolism.The common phenotype seen was a decrease in very-high-molecular-weight ubiquitin conjugate levels and the accumu-lation of several faster-migrating polyubiquitin species, al-though this phenotype was more subtle in rvs161 sur4 shp1 andrvs161 sur4 ufd1-1 cells. In some cases, the level of monoubiq-uitin was altered (rvs161 sur4 doa1, rvs161 sur4 doa1�C, andrvs161 sur4 cdc48-3 cells). Based on these results, we believethat the synthesis of monoubiquitin, its conversion to high-molecular-weight conjugates, and the subsequent turnover ofthese conjugates all are necessary for the sur4-dependent sup-pression of rvs161 endocytosis defects.

DISCUSSION

The rate of glucose transporter endocytosis is drasticallyreduced in rvs161 cells, and this defect correlates with star-vation-induced death under low-glucose conditions. Theseresults suggest that the glucose starvation defect is linked tothe improper regulation of hexose transporter traffickingand/or degradation. Viability under starvation is linked tothe endo domain of Rvs161, as endo� mutants are inviablewhen starved and are defective in endocytosing glucosetransporters from the membrane. In addition, the loss ofSUR4 suppresses the starvation defect and reinitiates glu-cose transporter endocytosis.

How might the reinitiation of endocytosis remediate starva-tion-induced death? One possibility is that the accurate endo-

TABLE 1. Glucose concentration-dependent appearance/disappearance of Hxt2-GFP from the plasma membranea

Strain

% of cells with plasma membrane-associated Hxt2 at glucose concn:

6% 0.05% 6%

Wild type 1 � 0.5 95 � 3 14 � 5rvs161 2 � 1 97 � 3 95 � 4rvs161 sur4 1 � 1 97 � 5 21 � 4rvs161 sur4 doa1 3 � 2 92 � 4 94 � 6rvs161 sur4 doa1F417D F434D 1 � 0.5 93 � 3 18 � 4rvs161 sur4 doa1�C 2 � 0.5 94 � 3 92 � 4rvs161 sur4 cdc48-3b 6 � 4 88 � 5 94 � 6rvs161 sur4 ufd1-1b 1 � 0.5 99 � 3 97 � 4rvs161 sur4 shp1 1 � 0.5 95 � 5 89 � 5

a Cells were sequentially shifted from 6% glucose to 0.05% glucose for 1 h,and then back to 6% glucose for 5 h. Values are the averages from threeindependent experiments.

b Cells were assayed at 30°C.

FIG. 8. Cell factors regulating ubiquitin levels are required for theloss of SUR4 to suppress the endocytosis defects of rvs161 cells. Cellswere grown to exponential phase in YEPD at 30°C. They then weresequentially shifted to medium containing the indicated sugar concen-trations for the indicated times. Hxt2-GFP localization was visualizedby live-cell fluorescence microscopy. (B) Cells were shifted to YEPcontaining 2% galactose to induce the expression of the dominant-negative Rsp5-1. WT, wild type; Glu, glucose; hr, hours.


cytosis/degradation of sugar transporters acts as a regulatorystep that is necessary to maintain proper stoichiometric trans-porter ratios, which are critical for growth under nutrient stressconditions; rvs161 mutants accumulate high-affinity transport-ers in response to glucose starvation, and they lack the abilityto endocytose/degrade low-affinity transporters. Glucose trans-porters in mammalian cells, specifically Glut1, exist in dimericand tetrameric states, but these oligomerizations are not es-sential for glucose uptake (48). A nonfunctional chimera con-sisting of yeast Hxt1 and Hxt4 transporters inhibits the func-tion of wild-type glucose transporters (67). Whether thischimera functions as a dominant-negative mutant, directly in-teracting with and inhibiting specific glucose transporters, hasnot been studied. Therefore, is maintaining proper ratios ofhomo- and heterodimeric transporters necessary for viabilityunder various growth conditions? With that said, we cannotrule out that the reinitiation of global endocytosis itself reme-

diates the starvation defect rather than the specific reinitiationof proper stoichiometric transporter ratios. rvs161 cells aredefective for growth under sulfur and nitrogen starvation con-ditions, and the loss of SUR4 remediates these growth defectsas well (15).

An alternative scenario is that rvs161 cells internalize andmetabolize too much glucose due to defects in endocytosis andthe constitutive accumulation of sugar transporters at theplasma membrane, which depletes cellular ATP stores. Thefirst step in glucose utilization after internalization is a phos-phorylation event by hexose kinase, resulting in glucose-6-phosphate production and shunting through the glycolyticpathway (27). rvs161 cells accumulate the high-affinity glucosetransporter Hxt2 and may accumulate others, such as Hxt6 andHxt7. The concentration of glucose (2.7 mM) under starvationconditions is within the Km ranges of Hxt6 and Hxt7 (Km 1to 2 mM) as well as those of Hxt2 (Km 1.5 mM) and Hxt4

FIG. 9. rvs161 cells hyperaccumulate monoubiquitin, and the loss of SUR4 suppresses this defect. Whole-cell lysates were resolved bySDS-PAGE, and monoubiquitin and high-molecular-weight polyubiquitin conjugates were visualized immunologically using anti-ubiquitin poly-clonal antibodies. The levels of various ubiquitin species were normalized against actin using densitometry analysis. The values are the averagesfrom three experiments � standard deviations.


(Km 10 mM) (34). Here again the general reinitiation ofendocytosis/degradation would act as a balance in conjunctionwith the rate of biosynthesis in order to maintain proper trans-porter ratios at the plasma membrane. Preliminary studiesexamining the rate of glucose binding and internalization inrvs161 cells thus far have been inconclusive (E. Swain and J. T.Nickels, unpublished data).

In addition to its role in endocytosis, Rvs161 is required foractin repolarization following osmotic salt stress. Wild-typecells depolarize actin following salt stress and repolarize aftera period of adaptation, whereas rvs161 mutant cells depolarizeactin but are unable to repolarize afterwards (1, 69). The lossof SUR4 suppresses the actin polarization defect of rvs161 cellsunder conditions that are semipermissive for viability (3.4%NaCl) (1). However, it does not suppress the actin polarizationdefect of mutant cells grown under glucose starvation condi-tions (26) or under high salt stress, which results in inviability(6% NaCl) (45a). Thus, how the loss of SUR4 suppresses rvsdefects cannot be explained solely through its effects on theactin cytoskeleton. Interestingly, the glucose starvation defectof the endo-deficient Rvs161 point mutants correlates withtheir ability to form an Rvs161-Rvs167 complex, as the rvs161R113K allele cannot bind Rvs167, as determined by two-hybridanalyses (P. McCourt, J. Morgan, and J. T. Nickels, unpub-lished data). These results seem reasonable, as the loss ofSUR4 is able to remediate defects of rvs161 rvs167 cells (15).Thus, suppression can occur in the absence of any Rvs161- orRvs167-associated multiprotein complexes.

The Cdc48 binding domain of Doa1 was required for sugartransporter endocytosis in rvs161 sur4 cells. Moreover, directroles for Cdc48-Shp1 and Cdc48-Ufd1 were demonstrated.Cdc48-Shp1 regulates several membrane fusion events, includ-ing nuclear envelope growth and reforming the ER and Golgiassembly after mitosis (46, 47, 64). During ER-dependent deg-radation, Cdc48-Ufd1 extracts ubiquitylated substrates frommembranes for their degradation. In yeast, Cdc48-Ufd1 mobi-lizes the transcription factors Spt23 and Mga2 from mem-branes prior to their moving to the nucleus (33, 59, 66). Basedon these and other observations, Meyer and Popp (46) suggestthat the fundamental activity of Cdc48 is the energy-dependentremoval of ubiquitylated proteins from membranes. Once re-moved, these proteins are free to be degraded or deubiquity-lated. How Cdc48 regulates sugar transporter endocytosisneeds to be explored; understanding how it helps in reinitiatingsphingolipid-dependent endocytosis should be an excellentmodel for study.

The level of monoubiquitin was increased in rvs161 cells, andthe deletion of SUR4 remediated this defect to some extent.Monoubiquitin serves as a regulatory signal for the intracellu-lar transport of proteins through the late secretory and endo-cytic pathways (reviewed in references 31 and 32). In yeast,membrane proteins such as amino acid permeases and matingfactor receptors are ubiquitylated, and this modification acts asa signal for internalization and/or endosomal sorting (reviewedin references 38 and 65). The mammalian sugar transportersGLUT1 and GLUT4 are modified with ubiquitin as well as theubiquitin-like protein SUMO (22, 41). Whether yeast glucosetransporters exhibit a similar fate is not known. However, thesphingolipid-dependent reinitiation of endocytosis has a strictrequirement for factors regulating ubiquitin pools, pointing to

the ubiquitylation of glucose transporters being an importantregulatory step in their turnover.

The N-BAR domain of the BAR family of proteins plays arole in initiating and/or sensing membrane curvature (58). Ifyeast Rvs161 and/or Rvs167 sense and are drawn to nascentbuds destined for endocytosis, they may act in the formationand function of a large membrane-associated multiprotein as-sembly complex that is involved in bud fission and subsequentendocytosis (21). Rvs167 does associate with the fission ma-chinery; however, this is a late event in the scission process(37). sur4 cells accumulate the sphingoid LCB phytosphin-gosine (51). The endogenous addition of this LCB reinitiatesendocytosis in cells deficient in sphingolipid biosynthesis (23,73), and its transient accumulation may be required for ubiq-uitin-dependent proteolysis following heat stress (11). LCBaccumulation may constitutively alter membrane curvature,causing invaginations within specific microdomains. These maygive rise to putative pseudonascent buds that normally areinitiated or stabilized by Rvs161-Rvs167 complexes, which thencan attract factors that are required for fission. An importantquestion remaining is whether LCB-dependent endocytosis re-quires the general fission machinery to initiate bud scission andrelease, as it does require ubiquitylation.

Defects in sphingolipid biosynthesis also could activate sig-naling pathways required to remediate rvs defects; activatingthese pathways would circumvent the need for any Rvs-depen-dent events during endocytosis. Sphingolipid intermediates inyeast, as in mammalian cells (16, 36, 52), are important signal-ing molecules, particularly the sphingoid base phytosphin-gosine. The accumulation of LCBs in yeast activates the Pkc1-MAP cell wall integrity pathway (74). The accumulation of themammalian LCB sphingosine 1-phosphate in yeast stimulatesCa2� accumulation, possibly initiating the LCB-dependent ac-tivation of Ca2�/calmodulin-dependent signaling cascades (5).

Whether the sphingolipid mutations suppressing rvs defectsall function by activating some LCB-dependent event(s) is notknown. sur4 mutants accumulate phytosphingosine but alsoaccumulate C22 fatty acids, are devoid of C26 fatty acids, pro-duce complex sphingolipids that have shorter fatty acid moi-eties, and hyperaccumulate an inositolphosphorylceramidespecies (51). Whether any of these changes in lipid metabolismcontribute to suppression is not known. Moreover, sur2 cells donot make phytosphingosine; thus, all complex sphingolipids arederived from the LCB dihydrosphingosine, and they lack C4

hydroxylation (29). Whether the loss of SUR2 and the accu-mulation of dihydrosphingosine activates pathways identical tothose regulated by phytosphingosine is not known.

A number of sphingolipid mutations suppressing rvs161 de-fects alter calcium homeostasis (75). Csg2 and Sur1 are re-quired for the synthesis of mannose inositolphosphorylcer-amide, and csg2 and sur1 mutants are calcium sensitive (3, 4).Recessive mutations in SUR2 and SUR4 remediate the calciumsensitivity of csg2 and sur1 cells (75). Interestingly, the additionof exogenous phytosphingosine alone to csg2 cells remediatestheir calcium-sensitive phenotype (75). We have been unsuc-cessful in suppressing rvs defects through calcium remediationor phytosphingosine supplementation. The endocytosis defectof rvs cells may preclude using exogenous methods; thus, moredetailed studies are warranted.


ACKNOWLEDGMENTS

We thank Howard Riezman and Michihiro Kasahara for anti-Suc2and anti-Gal2 antibodies, respectively. We are grateful to KeithWilkinson, James Mullally, Dale Haines, Valeria Brizzio, and MarkRose for strains. We thank Martin Adelson and Eli Mordechai formany helpful discussions.

This work was supported in part by NIH grant HL67401 (J.T.N.).We acknowledge and appreciate the financial support of Medical Di-agnostics Laboratories, L.L.C.

REFERENCES

1. Balguerie, A., M. Bagnat, M. Bonneu, M. Aigle, and A. M. Breton. 2002.Rvs161p and sphingolipids are required for actin repolarization followingsalt stress. Eukaryot. Cell 1:1021–1031.

2. Bauer, F., M. Urdaci, M. Aigle, and M. Crouzet. 1993. Alteration of a yeastSH3 protein leads to conditional viability with defects in cytoskeletal andbudding patterns. Mol. Cell. Biol. 13:5070–5084.

3. Beeler, T., K. Gable, C. Zhao, and T. Dunn. 1994. A novel protein, CSG2p,is required for Ca2� regulation in Saccharomyces cerevisiae. J. Biol. Chem.269:7279–7284.

4. Beeler, T. J., D. Fu, J. Rivera, E. Monaghan, K. Gable, and T. M. Dunn.1997. SUR1 (CSG1/BCL21), a gene necessary for growth of Saccharomycescerevisiae in the presence of high Ca2� concentrations at 37°C is requiredfor mannosylation of inositolphosphorylceramide. Mol. Gen. Genet. 255:570–579.

5. Birchwood, C. J., J. D. Saba, R. C. Dickson, and K. W. Cunningham. 2001.Calcium influx and signaling in yeast stimulated by intracellular sphingosine1-phosphate accumulation. J. Biol. Chem. 276:11712–11718.

6. Boles, E., and C. P. Hollenberg. 1997. The molecular genetics of hexosetransport in yeasts. FEMS Microbiol. Rev. 21:85–111.

7. Breton, A. M., J. Schaeffer, and M. Aigle. 2001. The yeast Rvs161 and Rvs167proteins are involved in secretory vesicles targeting the plasma membraneand in cell integrity. Yeast 18:1053–1068.

8. Brizzio, V., A. E. Gammie, and M. D. Rose. 1998. Rvs161p interacts withFus2p to promote cell fusion in Saccharomyces cerevisiae. J. Cell Biol.141:567–584.

9. Casal, E., L. Federici, W. Zhang, J. Fernandez-Recio, E. M. Priego, R. N.Miguel, J. B. DuHadaway, G. C. Prendergast, B. F. Luisi, and E. D. Laue.2006. The crystal structure of the BAR domain from human Bin1/amphiphy-sin II and its implications for molecular recognition. Biochemistry 45:12917–12928.

10. Chitu, V., and E. R. Stanley. 2007. Pombe Cdc15 homology (PCH) proteins:coordinators of membrane-cytoskeletal interactions. Trends Cell Biol. 17:145–156.

11. Chung, N., G. Jenkins, Y. A. Hannun, J. Heitman, and L. M. Obeid. 2000.Sphingolipids signal heat stress-induced ubiquitin-dependent proteolysis.J. Biol. Chem. 275:17229–17232.

12. Cleveland, D. W., S. G. Fischer, M. W. Kirschner, and U. K. Laemmli. 1977.Peptide mapping by limited proteolysis in sodium dodecyl sulfate and anal-ysis by gel electrophoresis. J. Biol. Chem. 252:1102–1106.

13. Crouzet, M., M. Urdaci, L. Dulau, and M. Aigle. 1991. Yeast mutant affectedfor viability upon nutrient starvation: characterization and cloning of theRVS161 gene. Yeast 7:727–743.

14. David, C., P. S. McPherson, O. Mundigl, and P. de Camilli. 1996. A role ofamphiphysin in synaptic vesicle endocytosis suggested by its binding to dy-namin in nerve terminals. Proc. Natl. Acad. Sci. USA 93:331–335.

15. Desfarges, L., P. Durrens, H. Juguelin, C. Cassagne, M. Bonneu, and M.Aigle. 1993. Yeast mutants affected in viability upon starvation have a mod-ified phospholipid composition. Yeast 9:267–277.

16. Dickson, R. C. 1998. Sphingolipid functions in Saccharomyces cerevisiae:comparison to mammals. Annu. Rev. Biochem. 67:27–48.

17. Dickson, R. C., and R. L. Lester. 2002. Sphingolipid functions in Saccharo-myces cerevisiae. Biochim. Biophys. Acta 1583:13–25.

18. Dickson, R. C., E. E. Nagiec, G. B. Wells, M. M. Nagiec, and R. L. Lester.1997. Synthesis of mannose-(inositol-P)2-ceramide, the major sphingolipidin Saccharomyces cerevisiae, requires the IPT1 (YDR072c) gene. J. Biol.Chem. 272:29620–29625.

19. Dickson, R. C., C. Sumanasekera, and R. L. Lester. 2006. Functions andmetabolism of sphingolipids in Saccharomyces cerevisiae. Prog. Lipid Res.45:447–465.

20. Durrens, P., E. Revardel, M. Bonneu, and M. Aigle. 1995. Evidence for abranched pathway in the polarized cell division of Saccharomyces cerevisiae.Curr. Genet. 27:213–216.

21. Engqvist-Goldstein, A. E., and D. G. Drubin. 2003. Actin assembly andendocytosis: from yeast to mammals. Annu. Rev. Cell Dev. Biol. 19:287–332.

22. Fernandes, R., A. L. Carvalho, A. Kumagai, R. Seica, K. Hosoya, T. Ter-asaki, J. Murta, P. Pereira, and C. Faro. 2004. Downregulation of retinalGLUT1 in diabetes by ubiquitinylation. Mol. Vis. 10:618–628.

23. Friant, S., R. Lombardi, T. Schmelzle, M. N. Hall, and H. Riezman. 2001.

Sphingoid base signaling via Pkh kinases is required for endocytosis in yeast.EMBO J. 20:6783–6792.

24. Gallop, J. L., and H. T. McMahon. 2005. BAR domains and membranecurvature: bringing your curves to the BAR. Biochem. Soc. Symp. 2005:223–231.

25. Gammie, A. E., V. Brizzio, and M. D. Rose. 1998. Distinct morphologicalphenotypes of cell fusion mutants. Mol. Biol. Cell 9:1395–1410.

26. Germann, M., E. Swain, L. Bergman, and J. T. Nickels, Jr. 2005. Charac-terizing the sphingolipid signaling pathway that remediates defects associ-ated with loss of the yeast amphiphysin-like orthologs, Rvs161p andRvs167p. J. Biol. Chem. 280:4270–4278.

27. Goncalves, P., and R. J. Planta. 1998. Starting up yeast glycolysis. TrendsMicrobiol. 6:314–319.

28. Grabs, D., V. I. Slepnev, Z. Songyang, C. David, M. Lynch, L. C. Cantley, andP. De Camilli. 1997. The SH3 domain of amphiphysin binds the proline-richdomain of dynamin at a single site that defines a new SH3 binding consensussequence. J. Biol. Chem. 272:13419–13425.

29. Haak, D., K. Gable, T. Beeler, and T. Dunn. 1997. Hydroxylation of Saccha-romyces cerevisiae ceramides requires Sur2p and Scs7p. J. Biol. Chem.272:29704–29710.

30. Habermann, B. 2004. The BAR-domain family of proteins: a case of bendingand binding? EMBO Rep. 5:250–255.

31. Hicke, L. 2001. Protein regulation by monoubiquitin. Nat. Rev. Mol. CellBiol. 2:195–201.

32. Hicke, L., and R. Dunn. 2003. Regulation of membrane protein transport byubiquitin and ubiquitin-binding proteins. Annu. Rev. Cell Dev. Biol. 19:141–172.

33. Hitchcock, A. L., H. Krebber, S. Frietze, A. Lin, M. Latterich, and P. A.Silver. 2001. The conserved npl4 protein complex mediates proteasome-dependent membrane-bound transcription factor activation. Mol. Biol. Cell12:3226–3241.

34. Ito, H., Y. Fukuda, K. Murata, and A. Kimura. 1983. Transformation ofintact yeast cells treated with alkali cations. J. Bacteriol. 153:163–168.

35. Itoh, T., and P. De Camilli. 2006. BAR, F-BAR (EFC) and ENTH/ANTHdomains in the regulation of membrane-cytosol interfaces and membranecurvature. Biochim. Biophys. Acta 1761:897–912.

36. Jenkins, G. M. 2003. The emerging role for sphingolipids in the eukaryoticheat shock response. Cell Mol. Life Sci. 60:701–710.

37. Kaksonen, M., C. P. Toret, and D. G. Drubin. 2005. A modular design for theclathrin- and actin-mediated endocytosis machinery. Cell 123:305–320.

38. Katzmann, D. J., G. Odorizzi, and S. D. Emr. 2002. Receptor downregula-tion and multivesicular-body sorting. Nat. Rev. Mol. Cell Biol. 3:893–905.

39. Kohlwein, S. D., S. Eder, C. S. Oh, C. E. Martin, K. Gable, D. Bacikova, andT. Dunn. 2001. Tsc13p is required for fatty acid elongation and localizes toa novel structure at the nuclear-vacuolar interface in Saccharomyces cerevi-siae. Mol. Cell. Biol. 21:109–125.

40. Krampe, S., O. Stamm, C. P. Hollenberg, and E. Boles. 1998. Cataboliteinactivation of the high-affinity hexose transporters Hxt6 and Hxt7 of Sac-charomyces cerevisiae occurs in the vacuole after internalization by endocy-tosis. FEBS Lett. 441:343–347.

41. Lalioti, V. S., S. Vergarajauregui, D. Pulido, and I. V. Sandoval. 2002. Theinsulin-sensitive glucose transporter, GLUT4, interacts physically with Daxx.Two proteins with capacity to bind Ubc9 and conjugated to SUMO1. J. Biol.Chem. 277:19783–19791.

42. Lichte, B., R. W. Veh, H. E. Meyer, and M. W. Kilimann. 1992. Amphiphysin,a novel protein associated with synaptic vesicles. EMBO J. 11:2521–2530.

43. Longtine, M. S., A. McKenzie III, D. J. Demarini, N. G. Shah, A. Wach, A.Brachat, P. Philippsen, and J. R. Pringle. 1998. Additional modules ofversatile and economical pcr-based gene deletion and modification in Sac-charomyces cerevisiae. Yeast 14:953–961.

44. Manolescu, A. R., K. Witkowska, A. Kinnaird, T. Cessford, and C. Cheese-man. 2007. Facilitated hexose transporters: new perspectives on form andfunction. Physiology (Bethesda) 22:234–240.

45. Masuda, M., S. Takeda, M. Sone, T. Ohki, H. Mori, Y. Kamioka, and N.Mochizuki. 2006. Endophilin BAR domain drives membrane curvature bytwo newly identified structure-based mechanisms. EMBO J. 25:2889–2897.

45a.McCourt, P., J. M. Morgan, and J. T. Nickels, Jr. Stress-induced ceramide-activated protein phosphatase can compensate for loss of amphiphysin-likeactivity in Saccharomyces cerevisiae and functions to reinitiate endocytosis.J. Biol. Chem., in press.

46. Meyer, H., and O. Popp. 2008. Role(s) of Cdc48/p97 in mitosis. Biochem.Soc. Trans. 36:126–130.

47. Meyer, H. H. 2005. Golgi reassembly after mitosis: the AAA family meets theubiquitin family. Biochim. Biophys. Acta 1744:481–492.

48. Mueckler, M. 1994. Facilitative glucose transporters. Eur. J. Biochem. 219:713–725.

49. Mullally, J. E., T. Chernova, and K. D. Wilkinson. 2006. Doa1 is a Cdc48adapter that possesses a novel ubiquitin binding domain. Mol. Cell. Biol.26:822–830.

50. Munn, A. L., B. J. Stevenson, M. I. Geli, and H. Riezman. 1995. end5, end6,and end7: mutations that cause actin delocalization and block the internal-


ization step of endocytosis in Saccharomyces cerevisiae. Mol. Biol. Cell6:1721–1742.

51. Oh, C. S., D. A. Toke, S. Mandala, and C. E. Martin. 1997. ELO2 and ELO3,homologues of the Saccharomyces cerevisiae ELO1 gene, function in fattyacid elongation and are required for sphingolipid formation. J. Biol. Chem.272:17376–17384.

52. Ohanian, J., and V. Ohanian. 2001. Sphingolipids in mammalian cell signal-ling. Cell Mol. Life Sci. 58:2053–2068.

53. Ozcan, S., J. Dover, and M. Johnston. 1998. Glucose sensing and signaling bytwo glucose receptors in the yeast Saccharomyces cerevisiae. EMBO J.17:2566–2573.

54. Ozcan, S., J. Dover, A. G. Rosenwald, S. Wolfl, and M. Johnston. 1996. Twoglucose transporters in Saccharomyces cerevisiae are glucose sensors thatgenerate a signal for induction of gene expression. Proc. Natl. Acad. Sci.USA 93:12428–12432.

55. Ozcan, S., and M. Johnston. 1995. Three different regulatory mechanismsenable yeast hexose transporter (HXT) genes to be induced by differentlevels of glucose. Mol. Cell. Biol. 15:1564–1572.

56. Ozcan, S., and M. Johnston. 1999. Function and regulation of yeast hexosetransporters. Microbiol. Mol. Biol. Rev. 63:554–569.

57. Ozcan, S., L. G. Vallier, J. S. Flick, M. Carlson, and M. Johnston. 1997.Expression of the SUC2 gene of Saccharomyces cerevisiae is induced by lowlevels of glucose. Yeast 13:127–137.

58. Peter, B. J., H. M. Kent, I. G. Mills, Y. Vallis, P. J. Butler, P. R. Evans, andH. T. McMahon. 2004. BAR domains as sensors of membrane curvature: theamphiphysin BAR structure. Science 303:495–499.

59. Rape, M., T. Hoppe, I. Gorr, M. Kalocay, H. Richly, and S. Jentsch. 2001.Mobilization of processed, membrane-tethered SPT23 transcription factorby CDC48(UFD1/NPL4), a ubiquitin-selective chaperone. Cell 107:667–677.

60. Reifenberger, E., E. Boles, and M. Ciriacy. 1997. Kinetic characterization ofindividual hexose transporters of Saccharomyces cerevisiae and their rela-tion to the triggering mechanisms of glucose repression. Eur. J. Biochem.245:324–333.

61. Ren, G., P. Vajjhala, J. S. Lee, B. Winsor, and A. L. Munn. 2006. The BARdomain proteins: molding membranes in fission, fusion, and phagy. Micro-biol. Mol. Biol. Rev. 70:37–120.

62. Ren, J., N. Pashkova, S. Winistorfer, and R. C. Piper. 2008. DOA1/UFD3plays a role in sorting ubiquitinated membrane proteins into multivesicularbodies. J. Biol. Chem. 283:21599–21611.

63. Revardel, E., M. Bonneau, P. Durrens, and M. Aigle. 1995. Characterization

of a new gene family developing pleiotropic phenotypes upon mutation inSaccharomyces cerevisiae. Biochim. Biophys. Acta 1263:261–265.

64. Romisch, K. 2006. Cdc48p is UBX-linked to ER ubiquitin ligases. TrendsBiochem. Sci. 31:24–25.

65. Rotin, D., O. Staub, and R. Haguenauer-Tsapis. 2000. Ubiquitination andendocytosis of plasma membrane proteins: role of Nedd4/Rsp5p family ofubiquitin-protein ligases. J. Membr. Biol. 176:1–17.

66. Shcherbik, N., and D. S. Haines. 2007. Cdc48p(Npl4p/Ufd1p) binds andsegregates membrane-anchored/tethered complexes via a polyubiquitin sig-nal present on the anchors. Mol. Cell 25:385–397.

67. Sherwood, P. W., I. Katic, P. Sanz, and M. Carlson. 2000. A glucose trans-porter chimera confers a dominant negative glucose starvation phenotype inSaccharomyces cerevisiae. Genetics 155:989–992.

68. Shupliakov, O., P. Low, D. Grabs, H. Gad, H. Chen, C. David, K. Takei, P.De Camilli, and L. Brodin. 1997. Synaptic vesicle endocytosis impaired bydisruption of dynamin-SH3 domain interactions. Science 276:259–263.

69. Sivadon, P., F. Bauer, M. Aigle, and M. Crouzet. 1995. Actin cytoskeletonand budding pattern are altered in the yeast rvs161 mutant: the Rvs161protein shares common domains with the brain protein amphiphysin. Mol.Gen. Genet. 246:485–495.

70. Sivadon, P., M. Crouzet, and M. Aigle. 1997. Functional assessment of theyeast Rvs161 and Rvs167 protein domains. FEBS Lett. 417:21–27.

71. Wang, Q., H. Y. Kaan, R. N. Hooda, S. L. Goh, and H. Sondermann. 2008.Structure and plasticity of Endophilin and Sorting Nexin 9. Structure 16:1574–1587.

72. Weissenhorn, W. 2005. Crystal structure of the endophilin-A1 BAR domain.J. Mol. Biol. 351:653–661.

73. Zanolari, B., S. Friant, K. Funato, C. Sutterlin, B. J. Stevenson, and H.Riezman. 2000. Sphingoid base synthesis requirement for endocytosis inSaccharomyces cerevisiae. EMBO J. 19:2824–2833.

74. Zhang, X., R. L. Lester, and R. C. Dickson. 2004. Pil1p and Lsp1p negativelyregulate the 3-phosphoinositide-dependent protein kinase-like kinase Pkh1pand downstream signaling pathways Pkc1p and Ypk1p. J. Biol. Chem. 279:22030–22038.

75. Zhao, C., T. Beeler, and T. Dunn. 1994. Suppressors of the Ca2�-sensitiveyeast mutant (csg2) identify genes involved in sphingolipid biosynthesis.Cloning and characterization of SCS1, a gene required for serine palmitoyl-transferase activity. J. Biol. Chem. 269:21480–21488.

76. Zhao, F. Q., and A. F. Keating. 2007. Functional properties and genomics ofglucose transporters. Curr. Genomics 8:113–128.


altering sphingolipid metabolism in saccharomyces ...cycling the plasma membrane at synaptic...

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